Mitochondria transplantation between living cells

Mitochondria and the complex endomembrane system are hallmarks of eukaryotic cells. To date, it has been difficult to manipulate organelle structures within single live cells. We developed a FluidFM-based approach to extract, inject, and transplant organelles from and into living cells with subcellular spatial resolution. The technology combines atomic force microscopy, optical microscopy, and nanofluidics to achieve force and volume control with real-time inspection. We developed dedicated probes that allow minimally invasive entry into cells and optimized fluid flow to extract specific organelles. When extracting single or a defined number of mitochondria, their morphology transforms into a pearls-on-a-string phenotype due to locally applied fluidic forces. We show that the induced transition is calcium independent and results in isolated, intact mitochondria. Upon cell-to-cell transplantation, the transferred mitochondria fuse to the host cells mitochondrial network. Transplantation of healthy and drug-impaired mitochondria into primary keratinocytes allowed monitoring of mitochondrial subpopulation rescue. Fusion with the mitochondrial network of recipient cells occurred 20 minutes after transplantation and continued for over 16 hours. After transfer of mitochondria and cell propagation over generations, donor mitochondrial DNA (mtDNA) was replicated in recipient cells without the need for selection pressure. The approach opens new prospects for the study of organelle physiology and homeostasis, but also for therapy, mechanobiology, and synthetic biology.


Introduction
Single-cell surgery approaches promise minimally invasive perturbation, i.e. removal or introduction of cellular compartments without compromising cell viability. Manipulation of mitochondria receives special emphasis due to their central cellular role: they are at the heart of energy conversion and link cellular metabolism to signaling pathways and cell fate decision (1-4). Mitochondria harbor their own genetic content (mtDNA), which is prone to accumulating erroneous, disease causing mutations (5)(6)(7) and are subject to quality control (8,9). Although mitochondria are generally inherited strictly vertically to daughter cells, exchange of larger cellular components including mitochondria has also been observed in tissues of multicellular organisms (10)(11)(12)(13). To reconstitute such transfer events, therapy approaches involve the grafting of purified mitochondria into a damaged area of a tissue or their intravenous injection (14). However the fate of these mitochondria is unknown (15).
With limited means to study and quantify the transfer of mitochondria into cells and without ways to analyze dose-response relationships experimentally, it is difficult to gain mechanistic insight on the actual impact of cytoplasmic and mitochondrial transmissions under healthy or diseased cellular states. Extraction and injection of organelles from and into single cells is technically demanding. Miniaturized probes have a high potential to manipulate and sample individual cells within their microenvironment at high spatiotemporal resolution (16).
Nano-scaled pipettes and nanotweezers allow sampling and trapping of individual charged molecules and single mitochondria (17,18) and have been combined with -omics methods, enabling compartment-resolved single cell studies (19,20). Other specialized microfluidic devices for microinjection into cultured cells have been introduced (21). Using a modified microcapillary pipet, mitochondria injection was achieved (22).
However, micropipette-based approaches are limited in terms of volume scalability, have only been applied to single mitochondria, and have been limited to either cell extraction or cell injection. In addition, the success rate of transferring mitochondria into single cultured cells has been low and requires use of cells artificially depleted of mtDNA with subsequent selection of transformed cells. This limits the approach to selective conditions and, in particular, it has prevented studies on the dynamic behavior of mitochondrial subpopulations to this point.
Despite all these crucial developments in single-cell technologies, functional transfer, i.e. transplantation of organelles from cell to cell, has not yet been achieved with exception of much larger oocytes.
FluidFM (23) combines the high-precision force-regulated approach of an atomic force microscope (AFM, pN to µN) with the volumetric dispensing of nano-scale pipets (fL to pL) under optical inspection, providing the forces and volume control relevant for single-cell manipulation (24). These features are unique among miniaturized probes and pivotal for driving the probe into the cytosolic compartment in a minimally invasive manner when delivering and extracting molecules (including plasmids, RNAs, and proteins) into-and from viable cells (25,26). In this study, we established FluidFM as a single-cell technology for intra-and intercellular micromanipulation of organelles in living eukaryotic cells (Fig. 1A-C). The fluidic handling of subcellular compartments poses a challenge for miniaturized probes because endomembrane structures are relatively large (mitochondria: 300-800 nm in diameter (27)) and form interconnected networks. Organelle manipulation also bears a higher risk of rupturing the cytoplasmic membrane compared to the few nm opening of probes used for molecule injection and extraction in prior work (25,26). Organelle extraction and injection achieved here required dedicated fabrication of tips with customized aperture area ( ), to both overcome steric constraints and increase the range of applicable suction forces ( ) at the aperture via the fluidic pressure ( ) in accordance to = × . We adapted the size and geometry of the aperture and microchannel of the FluidFM probes, and used the tips in combination with force control for membrane insertion via automated AFM and exertion of suction (i.e. tensile) forces via automated pressure controller.
We simultaneously inspected target cells and the transplant inside the transparent cantilever (via phase contrast and fluorescence microscopy, Figure 2A

Tunable organelle extraction from live cells
To enable organelle manipulations, in particular their unconstrained flow through the probe, we manufactured FluidFM-probes with a channel height of 1.7 µm and drilled apertures up to 1100 nm × 1100 nm ( = 1.2 µm 2 ) with focused ion beam (FIB) (Fig. 1B). In addition, we designed and fabricated dedicated probes with a cylindrical tip to facilitate minimally invasive cell entry. These were sharpened by FIB milling to resemble a hollow needle to facilitate membrane insertion (Fig. 1B, Supplementary Fig. 1). These probes had an aperture area of = 1.6 µm 2 , further minimizing steric limitations and increasing the range of applicable hydrodynamic forces spanning from a few pN to over 100 nN, while showing great robustness towards mechanical stress ( Supplementary Fig. 2). The general workflow for the manipulation of intracellular membrane enclosed compartments involves positioning the FluidFM probe above a selected subcellular location and their insertion by AFM force spectroscopy, followed by either extraction of material from the cell by exerting negative pressure (Fig. 1A, Supplementary Movie 1) or injection into the cell by positive pressure (Fig. 1C). Exclusion of large organelles is achieved by fine-tuning the aperture size ( ) and the strength of the applied negative pressure (− ). The extraction of cytoplasmic material is monitored in real time and the extract is inspected inside the FluidFM channel by optical microscopy after relieving the pressure ( = 0) and retracting the probe ( Fig. 2A-C, Supplementary Movie 1). FluidFM probes with a channel height of 1 µm can capture a total volume of 7 pL. The extraction process allows exclusion of organelle compartments that require larger apertures and higher hydrodynamic forces for extraction. However, when larger apertures are used, smaller and less strongly crosslinked organelles will be co-extracted (Fig. 1B).
The sampled material can be dispensed subsequently for downstream analyses or transplanted directly into a recipient cell (Fig. 1A). To examine the capabilities of the newly fabricated FluidFM probes for organelle sampling from single cells, we monitored the endoplasmic reticulum (ER) and mitochondria. We used human osteosarcoma epithelial (U2OS) cells and visualized in parallel the ER by expression of GFP fused to the resident protein Sec61β (sec61-GFP) and mitochondria by expression of BFP targeted to the mitochondrial matrix (su9-BFP). When utilizing pyramidal probes with an aperture size of = 0.5 µ 2 and low pressure offsets, < 20 mbar, we accumulated ER in the cantilever, which was accompanied by disappearance of GFP-signal in the cell (Fig. 2D, Supplementary Movie 2). During extraction, the ER was pulled towards the cantilever tip and we observed a general conversion of cisternal to tubular ER, in both U2OS cells and a similarly labelled kidney cell line (COS7, Supplementary Fig. 3, Supplementary Movies 2 and 3). Notably, under these conditions, the mitochondrial network remained unperturbed and mitochondria were not extracted.
Next, we aimed at extracting mitochondria using pyramidal probes with a larger aperture size ( = 1.2 µm 2 ) and newly developed, slanted cylindrical probes ( = 1.6 µm 2 ) ( We examined cell viability upon subcellular manipulation of ER and mitochondria and did not find it compromised (>95% cell viability) ( Supplementary Fig. 4). To further ensure that our extraction protocol does not damage the cytoplasmic membrane upon probe insertion, we conducted a dedicated set of experiments and monitored potentially occurring Ca 2+ influx from the cell culture medium using a fluorescent probe (mito-R-GECO1 (28)). Our experiments confirmed that there was no ion influx during and after manipulation, indicating integrity of the cytoplasmic membrane during organelle extraction (Supplementary Movies 7-8).
Monitoring mitochondrial extraction, we noticed that mitochondrial tubules exposed to tensile forces (negative pressure) underwent a shape transition reminiscent of a 'pearls-on-a-string phenotype' (29) inside the cytoplasm of the target cells. This phenotype was characterized by discrete spheres of mitochondrial matrix, connected by thin and elongated membrane stretches (Supplementary Fig. 5A-B). These globular structures eventually pinched off upon further exertion of a pulling force and resulted in spherical shaped mitochondria in the cantilever (Fig. 2E, Supplementary Figure 5A-C). To date, it was not possible to exert hydrodynamic forces intracellularly, while distinguishing physical manipulation from other potential cellular triggers. The observed mitochondrial 'pearls-on-a-string' phenotype was previously described to result from calcium overflow (29) or mitochondrial membrane rupture (30). To ensure mitochondrial membrane integrity and thus functionality, we investigated whether both mitochondrial membranes remained intact during the process. To

Mitochondrial transplantation into cultured cells
Our next goal was to demonstrate the functional delivery of mitochondria into new host cells and to achieve cell-to-cell organelle transplantation. In contrast to mitochondria extraction, for which both pyramidal probes and cylindrical probes could be used ( Supplementary Fig. 7). Quantification of the transplant showed that the number of transplanted mitochondria for these experiments varied from 3 to 15 mitochondria per cell (Fig. 3H). The different success rates between the two alternative protocols can be explained by differences in mitochondrial condition. When evaluating mitochondrial extraction protocols, we observed that a fraction of the extracted mitochondria undergo rupture of their outer membranes ( Supplementary Fig 8) (32). Irreversible damage of mitochondria leads to degradation inside cells and potentially cell apoptosis due to cytochrome c leakage. While cell-to-cell transplantation of mitochondria reduces throughput, it has the advantage that the extracellular time is short (< 1 min) and that mitochondria sampled by FluidFM are maximally concentrated in native cytoplasmic fluid, bypassing the use of artificial buffers altogether. We ensured that the extract remained near the aperture during extraction by filling the probes with immiscible perfluorooctane before extraction and transplantation.
Therefore, only small volumes (0.5 -2 pL) are injected into the host cells (Fig. 3B), up to the volume previously extracted from the donor cell (injection of larger volumes is automatically prevented due to inherent flow resistance properties of the pre-filled fluorocarbon liquid).
Labeling mitochondria of the recipient cell (su9-BFP) in addition to labeling donor cell mitochondria (su9-mCherry) allowed us to survey the state of the mitochondrial network in the transplanted cell. In both transfer approaches described above (transplantation and purification followed by injection), the tubular,

Fate of transplanted mitochondria in primary cells
Having developed an efficient protocol for cell-to-cell transplantation of mitochondria, we sought to test whether primary cells show similar uptake behavior as the tested cancer cells and, if so, what are the dynamics of integration of foreign mitochondria. We considered these particular experiments important because quality control mechanisms are impaired in cancer cell lines (33) and to demonstrate the broad applicability of the established protocol. In addition, several studies link naturally occurring mitochondrial transfer events with short-term benefits for individual cells and tissues, for example in osteocytes (34), adipose tissue (35) or in neurons (36). However, to the best of our knowledge, the fate of mitochondria or dose-response relationships have not be studied, and appropriate technologies of mitochondrial transfer that preserve cell viability have been lacking.
We used primary human endothelial keratinocytes (HEKa), a skin cell type that is generally susceptible to radiation damage and aging (37). In standard culture conditions, the mitochondrial network of HEKa cells is mostly tubular, forming a large connected network ( Supplementary Fig. 8) similar to HeLa cells studied above, indicating an active mitochondrial fusion machinery (38). In the experiments described above, the first fusion or degradation events occurred 20 minutes post transplantation and continued for more than 16 hours. Remarkably, the speed at which the primary transplant was processed was independent of transplant sizes. Fusion or degradation of large transplants, >40 mitochondria per cell, advanced at similar rate as smaller transplants, <8 mitochondria per cell (Supplementary Movie 12, Supplementary Fig. 11).
As outlined above, the established cell-to-cell transplantation protocol is minimally invasive with regard to the integrity of the mitochondria themselves when using 'healthy' donor cells, and cells receiving transplants from unperturbed donor cells showed mostly uptake of the transplant (Fig. 4H). Next, we wondered whether  Fig. 8). However, even upon H2O2 treatment , and even upon application of all drugs simultaneously to donor cells, mitochondrial acceptance in healthy recipient background was still comparable to all other conditions, indicating the potential of cells to cope with highly damaged mitochondria when occurring as isolated events (Fig. 4H, Supplementary Fig 12).
Since the tested conditions for drug-impaired mitochondria did not show an impact on mitochondrial uptake behavior, we wondered whether the amount of injected mitochondria might have an impact on transplantation outcome. Across all conditions, we transplanted 1117 mitochondria (Fig. 4I) and followed their fate in more than 100 individual primary cells after transplantation of 1 up to 53 mitochondria ( Supplementary Fig. 12).
Pooling uptake data of cross-tested conditions (n = 135

Mitochondria transplantation and transfer of mitochondrial genomes
After demonstrating the short-term response of cells to mitochondrial transplantation, we focused on the longterm effects of mitochondrial transfer over generations of host cells. Mitochondria differ from other membrane compartments in that they carry their own genome, which is propagated within the cell's inherent mitochondrial pool. It has been shown that mtDNA can be transferred into somatic cells via miniaturized probes under selective pressure, either by transfer into cells artificially rendered free of mitochondrial DNA (rho-zero cells) (22,44) (2-25% efficiency), or by selection using antibiotics (< 0.01% efficiency) (10). Therefore, the introduction of new mtDNA sequences into functional somatic cells remains a challenge (45,46). Before demonstrating the transfer of mtDNA into host cells upon transplantation, we first wondered whether the FluidFM-extracted mitochondria contain mitochondrial DNA, which is organized in discrete complexes termed mitochondrial nucleoids, because the extraction process leads to rapid fragmentation of the network.
To visualize the behavior of mitochondrial nucleoids during mitochondrial extraction using FluidFM, we expressed a fluorescently tagged version of p55 (p55-GFP), a polymerase-γ subunit that co-localizes with mitochondrial nucleoids and appears in discrete speckles scattered throughout the mitochondrial matrix (47) (Fig. 5A). We evaluated time-lapse experiments upon mitochondrial extraction with labelled nucleoids and counted mitochondrial fragments either containing labelled nucleoids, or being devoid of their fluorescent signal ( Fig. 5B and Supplementary Fig. 13). We observed p55-GFP in > 90% of formed mitochondrial spheres (n = 18), indicating that most transplanted mitochondria contained mtDNA.
Next, we applied genomics to quantify uptake and maintenance of mtDNA after mitochondria transplantation (Fig. 5C). We compared two methods of mitochondrial transfer, using U2OS cells as mitochondria donors and

Discussion
The technology developed here allows the manipulation of intracellular compartments using FluidFM. We show the removal and injection of organelles from and into single cells without compromising organelle integrity nor cell viability. The scalability of the volume of the subcellular sampling protocol for organelles allows molecular downstream analyses, which are increasingly feasible thanks to the improving sensitivity of 'omics' technologies (52). Samplings can be performed at one time point, but also repeatedly from one cell to unravel the dynamic behavior within individual cells. We demonstrate the application of FluidFM to exert localized fluidic forces within single cells and thus expand the possibilities for subcellular sampling as well as the study of organelle mechanobiology (17,26). It has been proposed that mechanical forces and membrane constriction affect mitochondrial shape and dynamics (53,54), but with the previously established tools it has been difficult to test such a hypothesis without perturbing the cellular state as a whole (30,53). Gonzalez-Rodriguez et al (30) suggest that mitochondrial shape depends on the elastocapillary number ( ) (55) describing the ratio between the elastic modulus ( ) and mitochondrial membrane tension ( ): ≡ × ⁄ where d is the membrane thickness. A decreasing results in 'pearling' (30,55). This description explains our observations where pulling on the mitochondrial membrane ( Supplementary Fig. 5A) increases the mitochondrial membrane tension, thus decreasing . While the application of compressive force can be controlled in time and space (56), application of controlled tensile force has been impossible to date. Here, we demonstrate that mechanical force can be a driver for mitochondrial shape transition that is strictly localized to sites of tensile force application and propagates along membrane-connected mitochondrial tubes. Here, the purely mechanical nature of FluidFM presents itself as a strength, because it allows dissection from complex physiological stimuli, often involving calcium-influx, from isolated exertion of (hydrodynamic) pulling forces.
'Pearling' eventually leads to recruitment of Drp1 and mitochondrial fission. In a more physiological setting, the transition into the pearls-on-a-string phenotype appears to be an elegant solution that protects cells against membrane leakage during mechanical stress.
Transplantation of mitochondria at a high efficiency allowed us to track organelle fate over time in new genetic and physiological cell backgrounds. Similar to organ transplants that are accepted or rejected by new hosts, here we show rescue or failure of organelles within single cells after transplantation. We show that transplantation is highly efficient (95% success rate) when mitochondria are transplanted directly between living cells rather than using isolation protocols prior to transfer. The FluidFM-based approach of efficient mitochondria transplantation permitted us to evaluate mitochondrial quality control in primary cells by transplanting healthy and compromised mitochondria and observing their fate. We show that individual cells generally differentiate between individual mitochondria, but do not display apparent responses to mitochondria previously exposed to the tested stresses. Across conditions, we observe that the majority of mitochondria become integrated into the host network and that transplanted mitochondria give rise to secondary mitochondrial particles. Such particles are reminiscent of mitochondrial-derived vesicles that take part in mitochondrial quality control (57,58). This indicates that the extent of mitochondrial quality control may depend on the general cellular state rather than the actual quality of the mitochondrial network. The study of mitochondrial quality control is of great interest, and the approach introduced here has the potential to significantly contribute further to this field by allowing defective mitochondria to be introduced locally in an otherwise functional cellular background. In addition, it will be interesting to study the impact of mitochondrial In the future, the technique introduced here will stimulate applications in additional research areas, for example, the rejuvenation of cells with low metabolic activity in stem cell therapies (4,5) or as an alternative strategy in mitochondrial replacement therapy approaches. Beyond, it offers new perspectives to address fundamental questions in cell biology, mechanobiology and cell engineering. Then, the probes were glued onto a cytoclip holder by Cytosurge. Before each experiment, the cantilevers were cleaned by a 90 s plasma treatment (Plasma Cleaner PDG-32G, Harrick Plasma) before coating overnight with vapor phase SL2 Sigmacote (Sigma-Aldrich) in a vacuum desiccator. The siliconized probe was oven dried at 100 °C for 1 h. The cantilever spring constant was measured using software-implemented scripts (cylindrical probes: 2 ± 0.4 Nm -1 , pyramidal probes: 5 ± 1 Nm -1 ).

FluidFM setup and Microscopy
The FluidFM setup is composed of a FlexAFM 5-NIR scan head controlled by a C3000 controller (Nanosurf), a digital pressure controller (ranging from -800 mbar to +1000 mbar), and Microfluidic Probes (Cytosurge).
The scan head is mounted on an inverted AxioObserver microscope equipped with a temperature-controlled incubation chamber (Zeiss). The microscope is coupled to a spinning disc confocal microscope (Visitron) with a Yokogawa CSU-W1 scan head and an EMCCD camera system (Andor). For all images and videos, a 63× oil objective with 1.4 numerical aperture and a 2× lens switcher was used (without lens switcher: 4.85 pixel/micron and 9.69 pixel per micron with lens switcher); images are in 16 bit format. Image acquisition was controlled using the VisiView software (Visitron); linear adjustments and video editing were made with Fiji(66); additionally, images and videos were noise-filtered using the wiener noise filtering function (wiener2; 3 by 3 neighborhood size) in MatlabR2018a (MathWorks). Movies were created using a self-written Matlab script in order to visualize several sections or channels within the same movie. Colormaps originate from Thyng et al. (67). Images of cantilevers containing extracts (Fig. 2) were created summing up the slices of a Zstack via Fiji and reconverting the image to 16 bit format.
For the experiments, the culture media was replaced with CO2-independent growth medium containing 10% FBS (ThermoFisher) and 1% penicillin-streptomycin (ThermoFisher). Cell lines stably expressing fluorescent proteins markers were created via lentiviral transduction, the constructs were previously described in Helle et al. (53).

Mitochondrial pulling and extraction experiments.
All experiments were executed at 37°C. The cells for extraction/transplantation were selected by light microscopy. Z-stacks were taken before and after the manipulation step to document the workflow. FluidFM probes were prepared as specified above and prefilled with octadecafluorooctane (perfluorooctane) (Sigma-Aldrich). Subsequently, the FluidFM probe was moved over a targeted area in the cytosol of a selected cell, usually close to the nucleus or mitochondrial tubes in the cell periphery. The cantilever was then inserted at the specified location driven by a forward force spectroscopy routine in contact mode, until the setpoint of 400 nanoNewtons (nN) was reached. The probe was then kept at this position (in the X-Y dimension) at the given force offset. Then, negative pressure in the range between -10 to -150 mbar was applied to aspirate cellular content. Before retracting the probe at the end of the aspiration process, the pressure was set back to 0 mbar. The force setpoint was adjusted by analyzing force distance curves from neighboring cells within the same experiment; the force value at which the curve takes a linear shape (in this case 80 nN) was estimated. This force value marks the point at which the probe makes contact with the glass bottom below the cell; consequently, this value was chosen as a setpoint for the extraction.

Extraction of the ER fraction:
The experiments were performed using pyramidal cantilevers featuring an aperture area of 0.5 µm 2 (see For probe insertion with minimal Ca 2+ influx as shown in Movie S7, newly coated (Sigmacote, see above) FluidFM cantilevers were used. To deliberately disturb the cell membrane (Movie S9), the probe was driven into the cell using the same setpoint, then the optical table (Newport) was gently flicked with the index finger to cause the probe to shift slightly, effectively disturbing the cell membrane. Cell viability was controlled using the LIVE/DEAD cell imaging kit (ThermoFisher).

Mitochondrial transplantation experiments. FluidFM injection of bulk-purified mitochondria:
Mitochondria were purified from approximately 2*10 6 cultured HeLa cells continuously expressing the mitochondrial matrix marker su9-mCherry using the Qproteome Mitochondria Isolation Kit (Qiagen) following manufacturer's instructions. After purification, the mitochondria were washed two times in injection buffer (see above), and finally resuspended in 40 µl injection buffer before being loaded into FluidFM-probes having a sharpened cylindrical apex.
Coupling mitochondrial extraction with transplantation from individual cell to cell: Mitochondria were aspired as described above, using FluidFM-probes having a sharpened cylindrical apex from HeLa-cells that were co-cultured on the same dish, previously seeded within another quadrant of the 4-well micro insets.
Subsequently, the probe was moved to a region containing cells targeted for transplantation. For injection of mitochondria, the probe was positioned above a target cell as described above for the extraction of organelles.
The cantilever containing the organelles was then inserted into the cell in contact mode (setpoint 100 nN). The   Table S1).

Analysis of mitochondrial quality control mechanisms with mitochondrial transplantation in primary
HEKa cells. Cells were seeded into as described above, into low µ-dishes (ibidi) inside two-well culture inserts (ibidi). The special separation allows for drug-treatments of the donor cell population before the experiment, while the host cell population of HEKa cells remains under standard culture conditions. At the beginning of the experiment, the cell media was replaced with CO2-independent growth medium containing 10% FBS (ThermoFisher) and 1% penicillin-streptomycin (ThermoFisher

Data availability
Data to understand and assess the conclusion of this research are available in the main text, raw data sets will be made available upon request.

Code availability
Matlab code used for image analysis and mitochondrial particle distribution will be made available upon request.